BAC Recombineering using the modified DH10B strain SW102 galK

Recombineering protocol #3
BAC Recombineering using the modified DH10B strain SW102
and a galK positive/counterselection cassette
by Søren Warming, Ph.D.
This protocol explains in detail how to use the galK positive and counterselection
scheme to make essentially any BAC modification (e.g. point mutations,
deletions, and insertions). The modified BAC can then be used directly for
making BAC transgenic mice by pro-nuclear microinjection, or a fragment
containing the modification can be retrieved using gap repair and turned into a
gene targeting vector for use in mouse ES cells. The BAC modifications are done
using a modified bacterial strain, SW102. This strain is derived from DY380 and
therefore contains the l prophage recombineering system. Furthermore, the
galactose operon has been modified so it is fully functional, except the
galactokinase gene (galK) has been deleted. Importantly, the galK function can
be added in trans, and thereby the ability to grow on galactose as carbon source
is restored. The galK selection scheme is a two-step system: First the galK
cassette, containing homology to a specified position in a BAC, is inserted, by
homologous recombination, into the BAC. The recombinant bacteria will now be
able to grow on minimal media with galactose as the only carbon source, so the
first step is positive selection. Second, the galK cassette is substituted by an
oligo (double- or singlestranded), a PCR product, or a cloned fragment with
homology flanking the cassette. This is achieved by selecting against the galK
cassette by resistance to 2-deoxy-galactose (DOG) on minimal plates with
glycerol as the carbon source. DOG is harmless, unless phosphorylated by
functional galK. Phosphorylation by galK turns DOG into 2-deoxy-galactose-1phosphate, a non-metabolizable and therefore toxic intermediate. From the
resulting DOG-resistant colonies some will be background colonies, where the
bacteria have lost the galK cassette by a deletion, and the rest will be truly
recombinant clones. Even with as low as 33 bp homology on both sides of the
oligo we have still observed an efficiency of 50% correct clones, and with 50 bp
homology arms we routinely observe 60-80% efficiency. Longer homology arms
might improve the efficiency even more. Chloramphenicol selection is used
throughout, in order to maintain the BAC.
1. Design galK primers with 50 bp homology to an area flanking the desired
site to be modified. The 3’ end of these primers bind to the galK cassette.
For example, if you want to make a single bp substitution, your homology
arms should be the 50 bp on either side of the single bp. This will result in
a deletion of that basepair in the first step. If you want to make a small or a
large deletion, design the galK primers so the deletion is made already in
the first step. The primers should look as follows:
2. Design other primers/oligos depending on what the desired modification
is. If point mutation, design two 100 bp complementary oligos with the
modified basepair(s) in the middle. The remaining bases on either side are
homology arms. If a deletion is made with the galK cassette in the first
step, design two complementary oligos to substitute for galK. This will
result in a “seam-less” deletion, leaving nothing but the desired change
3. Choose a BAC for your purpose. If you need a 129 BAC for subsequent
construction of a targeting vector, screen a 129 library like the CITB library
from Invitrogen. Order the clones from Invitrogen. End sequence the
positive clones to find one that contains the sequence needed. If a
C57BL/6 BAC is needed, identify a BAC using a genome browser (like, and order the BAC from Invitrogen or CHORI.
Before proceeding, make sure the BAC is 100% correct; PCR analysis
and restriction enzymatic analysis (fingerprinting). We have had good
succes with SpeI digests – it cuts frequently enough to give many bands,
and infrequently enough so that the band sizes are informative. It’s
worthwhile spending some time on this BAC characterization – if you
don’t, you might regret it later…
4. Transform the characterized BAC into electrocompetent SW102 cells (see
recombineering protocol #1). Recover for 1 hour at 32°C, and plate on LB
plates with 12.5 mg/ml chloramphenicol.
5. PCR amplify the galK cassette using the primers designed in step 1 and a
proof-reading Taq-mix (we use Expand High Fidelity from Roche). Use 1-2
ng template (the pgalK plasmid). 94°C 15 sec., 60°C 30 sec., 72°C 1 min.,
for 30 cycles. Add 1-2 ml DpnI per 25 ml reaction, mix, and incubate at
37°C for 1 hour. This step serves to remove any plasmid template;
plasmid is methylated, PCR products are not. Gel-purify the DpnI-digested
PCR product, preferably overnight at low voltage. From a strong PCR
band, purified, and eluted in 50 ml ddH2O, we use 2.5 ml for a
transformation (approx. 10-30 ng).
6. Inoculate an overnight culture of SW102 cells containing the BAC. 5 ml LB
+ chloramphenicol. Incubate at 32°C.
7. Next day, turn on two shaking waterbaths: One at 32°C, the other at 42°C.
Make an ice/water slurry and put a 50 ml tube of ddH2O in there to make
sure it’s ice-cold (see later). Dilute 500 ml of the overnight SW102 culture
containing the target BAC in 25 ml LB with chloramphenicol (12.5 mg/ml) in
a 50 ml baffled conical flask and incubate at 32°C in a shaking waterbath
to an OD600 of approx. 0.6 (0.55-0.6). This usually takes 3-4 hours.
8. Transfer 10 ml to another baffled 50 ml conical flask and heat-shock at
42°C for exactly 15 min. in a shaking waterbath. The remaining culture is
left at 32°C as the un-induced control.
9. After 15 min., the two samples, induced and un-induced, are briefly cooled
in an ice/waterbath slurry and then transferred to two 15 ml Falcon tubes
and pelleted using 5000 RPM at 0°C for 5 min. It’s important to keep the
bacteria as close to 0°C as possible in order to get good competents cells.
10. Pour off all of the supernatant and resuspend the pellet in 1 ml ice-cold
ddH2O by gently swirling the tubes in the ice/waterbath slurry. No
pipetting. This step may take a while. When resuspended, add another 9
ml ice-cold ddH2O and pellet the samples again.
11. Repeat step 10.
12. After the second washing and centrifugation step, all supernatant must be
removed by inverting the tubes on a paper towel, and the pellet
(approximately 50 ml each) is kept on ice until electroporated with PCR
13. Transform the now electrocompetent SW102 cells. We use 25 ml cells for
each electroporation in a 0.1 cm cuvette (BioRad) at 25 mF, 1.75 kV, and
200 ohms. After electroporation of the PCR product, the bacteria are
recovered in 1 ml LB (15 ml Falcon tube) for 1 hour in a 32°C shaking
14. After the recovery period the bacteria are washed twice in 1xM9 salts (see
appendix A) as follows: 1 ml culture is pelleted in an eppendorf tube at
13,200 RPM for 15 sec. and the supernatant removed with a pipette. The
pellet is resuspended in 1 ml 1xM9 salts, and pelleted again. This washing
step is repeated once more. After the second wash, the supernatant is
removed and the pellet is resuspended in 1 ml 1xM9 salts before plating
serial dilutions in 1xM9 (100 ml, 100 ml of a 1:10 dilution, and 100 ml 1:100)
onto M63 minimal media plates (see appendix A) with galactose, leucine,
biotin, and chloramphenicol. Washing in M9 salts is necessary to remove
any rich media from the bacteria prior to selection on minimal media. The
uninduced samples routinely have a higher degree of lysis/bacterial death
after electroporation and you will lose some bacteria, so the uninduced
sample is diluted in 0.25 – 0.75 ml 1xM9 salts in the final step to make up
for the difference. Plate 100 ml of the uninduced sample as a control.
15. Incubate 3 days at 32°C in a cabinet-type incubator.
16. Streak a few colonies onto MacConkey + galactose + chloramphenicol
indicator plates. Streak to obtain single colonies (see appendix B). The
colonies appearing after the 3 days of incubation should be Gal+, but in
order to get rid of any Gal- contaminants (hitch-hikers), it is important to
obtain single, bright red colonies before proceeding to the second step.
Gal- colonies will be white/colorless and the Gal+ bacteria will be bright
red/pink due to a pH change resulting from fermented galactose after an
overnight incubation at 32°C.
17. Pick a single, bright red (Gal+) colony and inoculate a 5 ml LB +
chloramphenicol overnight culture. Incubate at 32°C. There is normally no
need to further characterize the clones after the first step.
18. Repeat steps 7 through 12 above to obtain electrocompetent SW102 cells
(now ready for a galK <> mutation substitution). If you are going to
transform a double-stranded DNA oligo, the two complementary oligos
can be annealed in vitro: Mix 10 mg of each oligo in a volume of 100 ml 1x
PCR buffer. Boil for 5 min. Let cool slowly to room temp (30 min.). Add 10
ml 3 M NaAc and 250 ml EtOH. Precipitate, wash once in 70% EtOH, and
resuspend the final, air-dried, pellet in 100 ml ddH2O (final conc. of 200
ng/ml). Use 1 ml per transformation.
19. Transform the bacteria (25 ml of heat-shocked and 25 ml of uninduced
control) with 200 ng double-stranded oligo, a PCR product, or anything
containing a mutation and with homology to the area flanking the galK
cassette. Recover in 10 ml LB in a 50 ml baffled conical flask by
incubating in a 32°C shaking waterbath for 4.5 hours. This long recovery
period serves to obtain bacteria, by “dilution”, that only contains the
desired recombined BAC, and thus have lost any BAC still containing the
galK cassette.
20. As in step 14, pellet 1 ml culture and wash twice in 1xM9 salts, and
resuspend in 1 ml 1xM9 salts after the second wash before plating serial
dilutions (100 ml, 100 ml of a 1:10 dilution, 100 ml 1:100, and 100 ml 1:1000)
on M63 minimal media plates with glycerol, leucine, biotin, 2-deoxygalactose (DOG), and chloramphenicol.
21. Incubate at 32°C for three days.
22. The number of colonies may or may not be significantly different when
comparing plates from uninduced and induced bacteria (range between
1:1 – 1:100). In either case, you will still be able to find true recombinants
with a high frequency. Analyze, say, 10-12 colonies by SpeI digestion of
BAC miniprep DNA (see appendix C). Include a SpeI digest of the parent
BAC as a control. Clones with a digestion pattern like the parent are likely
to have undergone the desired mutation. Background clones (DOG
resistant without the desired mutation) willl have obvious deletions, and
should not be analyzed further. The clones with correct digestion pattern
should be analyzed by PCR and sequencing of the mutated region. If the
pattern is identical to the parental digestion pattern, it means that the
bacteria most likely became DOG resistant due to the desired homologous
recombination event. Alternatively, any deletion that contains the region
with the galK cassette, but excludes the chloramphenicol region, will also
be selected in the counterselection procedure. With a high frequency of
homologous recombination as in he SW102 strain, the background is not
likely to be a problem. In the unlikely event that too high a background is
observed, try to increase the length of the homology arms, to increase the
frequency of homologous recombination.
Appendix A. Media (from Current Protocols in Molecular Biology)
M9 medium (1 liter)
6 g Na2HPO4
3 g KH2PO4
1 g NH4Cl
0.5 g NaCl
M63 minimal plates
1L 5X M63
10 g (NH4)2SO4
68 g KH2PO4
2.5 mg FeSO4·7H2O
adjust to pH 7 with KOH
0.2 mg/ml d-biotin (sterile filtered) (1:5000)
20% galactose (autoclaved) (1:100)
20% 2-deoxy-galactose (autoclaved) (1:100)
20% glycerol (autoclaved) (1:100)
10 mg/ml L-leucine (1%, heated, then cooled down and
sterile filtered)
25 mg/ml Chloramphenicol in EtOH (1:2000)
1 M MgSO4·7H2O (1:1000)
Autoclave 15 g agar in 800 ml H2O in a 2 liter flask. Let cool down a little. Add
200 ml autoclaved 5X M63 medium and 1 ml 1 M MgSO4·7H2O. Adjust
volume to 1 liter with H2O if necessary. Let cool down to 50°C (“touchable
hot”). Add 10 ml carbon source (final conc. 0.2%), 5 ml biotin (1 mg), 4.5 ml
leucine (45 mg), and 500 ml Chloramphenicol (final conc. 12.5 mg/ml). Pour
the plates, 25-40 plates per liter.
MacConkey indicator plates
Prepare MacConkey agar plus galactose according to manufacturer’s
instructions. After autoclaving and cooling to 50°C, to one liter add 500 ml
Chloramphenicol (final conc. 12.5 mg/ml), and pour the plates, 25-40 plates
per liter.
Appendix B. Streaking for single colonies.
Refer to the figure below (from Current Protocols in Molecular Biology). With
a sterile toothpick, streak a single colony on a plate, in a forward-and-back
motion a couple of times. With a new toothpick, streak in a pattern
perpendicular to the first pattern, starting by streaking through the first
pattern. Repeat at least once more. This will result in a dilution of the bacterial
density, and you should obtain single colonies.
Appendix C. BAC minipreps.
For BAC minipreps (1-1.5 mg) we use the following protocol: 5 ml overnight
LB culture with chloramphenicol (15 ml Falcon tube) is pelleted for 5 min. at
5,000 RPM, the supernatant removed, and the pellet dissolved in 250 ml
buffer P1 (miniprep kit, Qiagen) and transferred to an eppendorf tube. 250 ml
P2 buffer is added, followed by mixing by inversion and incubation for <5 min.
at room temperature. Add 250 ml N3 buffer, followed by mixing and incubation
on ice for 5 min. The supernatant is cleared by two rounds of centrifugation at
13,200 RPM for 5 min. in a tabletop centrifuge. Each time the supernatant is
transferred to a new tube. DNA is precipitated by adding 750 ml isopropanol,
mixing and incubating on ice for 10 min., and centrifugation for 10 min. at
13,200 RPM. The pellet is washed once in 70% ethanol and the airdried pellet
is dissolved in 50 ml TE. 40 ml (approximately 1 mg) can be used for restriction
analysis in a 50 ml reaction, and 1 ml can be used as template for PCR
analysis or for transformation of electrocompetent bacteria.